His tag protein purification problem - (Jun/12/2004 )

I have seen that people changed the pH of the column according to the pI of their protein. Is there any general rule of thumb? The estimated pI of my 16kDa peptide is 9.38.

Cheers

Hi People!,

I was given two plasmids containing 2 HIS tagged proteins. I have the plasmids in BL21 cells and in DH5alfa cells.
I did a PCR to confirm the gene was there. I was also told that a secuence was made and it was allright. Before you ask I checked all the phs of the solutions myself, and they are fine.

The protein A is a pentamer that weights 28kda, isoelectric point 7,20, charge at ph7= 0,29. T he protein B is another pentamer that mw: 38Kda, isoelectric point 8,12, charge at ph 7, 1.28. (all calculations were made including the tag).

I grew 100ml of bacteria until OD=0,6, then I induced 1, 2, 3,4 hours with 1mM IPTG at 37C. I didn't use DTT. All was done at ambient temperature. I tried only the native step, I didn't denature the protein.

My coomassies aren't the greatest thing in the world, but they show *little* induction after 4 hours, almost none!, I'm supposed to see a super band <------------ Even more at 4 ours, I see like it's all degraded!

I see the protein in the dots in almost all the steps but I don't get to see the proteins in the western blot, or when I see it, it's a very light and timid band!.

To run the colum I use one wash without imidazole, then 10mMImidazole, 10mM Imidazole, 125, 125, 250, 500mM. The problem is that I see lots of bands in the elution step in the coomassie.

I use this parameters because the person that worked before with this proteins told me that that was the best, so I still don't change them. I use the recommended buffers in the QIAexpressionist book:
Lysis buffer, 500mM NaH2PO4, 300mMNaCl, 10mM Imidazole, ph8
Wash buffer: 50mM NaH2PO4, 300mM NaCl, ph8, Elution idem but with Imidazole

My Ab was against the protein and it wasn't very good, so today I made a western against the HIS tag (of the 1hr, 2hr, 3hr, 4hr extract), and the only thing that showed up was the RPN800 colored marked (no clue why this happened, could it be that the proteins of the marked are purified with HIS tags?). I only did one but I plan to start anti HIS westerns with ECL tomorrow.

What do you guys recommend me??
I would like to start with the induction process which is clearly not working well.
How high can the IPTG concentration go?

I use a 6x sample buffer with 0,5M of DTT.

I worked with all kinds of proteins with His-tag on N- and C- termini. I never had the problem you described here. If I was you I would have checked the DNA sequence first. Sometimes His-tag just does not get attached to the protein due to reading frame shift or insertion a stop codon before the His-tag.
I personally never got 100% pure protein using just Ni-NTA resin. E. coli contains some histidine-rich proteins with high affinity to Ni-NTA resin. It is absolutely normal to get 70-80% purity after this colomn.

Hi,

The plasmids are: PrSetA, in BL21 cells, and PQE70 in DH5alfa cells.
I know that DH5alfa are not the best thing to express protein, but shouldn't I see at least "some" induction?.
The person who worked before with these cells assured me the method worked, althought I never saw a coomassie.

Which other option could I try with IPTG to rule out the posibility that I'm not making the induction part wrong?

Best regards,
Carla

I've been trying on and off to establish a protocol to express and purify small proteins by Ni affinity chromatography. The last step of purifying my protein of interest away after Tev digest is just killing me and suggestions are welcomed. I must be doing something incredibly bone-headed.

I'm using Novagen's pet43a which has NusA fusion protein and 6XHis tag. During cloning, I engineered in a Tev cleaveage site between the tags and my coding region of interest. Expression in BL21(DE3) is no problemo. The fusion protein itself is 65kDa and my proteins of interest are 8-10 kDa.

Initial purification scheme #1(native conditions): lysis in 20mM sodium phosphate pH 7.4, 0.5 M NaCl and 10mM Imidazole. Purify on HiTrap chelating column from Amersham by eluting in 500mM Imidazole. Got a lot of fusion protein in fractions.

Desalt and digest with Tev enzyme(Tried both Invitrogen and homemade batch from colleague) at 30 degrees or 4 degrees. On SDS gel, digest looks like it has gone to completion. With a vivid imagination, sometimes a small molecular weight band bigger than my protein but smaller than fusion protein can be perceived The fusion protein is 75kDa and my cleaved protein of interest is 8.2kDa and so on a 17% gel, band migration might be kind of wonky and not be best indicator for size.

I've tried purification by anion exchange (Q XL with 20mM Tris+2M NaCl gradient), reverse phase, stir cell and got a whole of nothing, nothing, nothing.

So I figured, degradation is going on, right? So I try purification scheme #2 under denaturing conditions.

Lysis in 0.1M Tris pH8, 6M GdnHCl by stirring at room temp for 30-60 minutes. Centrifuged to clarify supernatant and purify it on HiTrap Chelating column. I got less protein in eluate under denaturing conditions than under native conditions which is weird; one would think the lack of secondary structure could enhance binding to the Ni column.

So I try a gravity drip column with the Amersham chelating sepharose slurry, maybe 1ml/min was too fast for binding kinetics. Eluted in 20mM sodium phosphate+500mM Imidazole and 8M urea and eluted fractions were not as impressive as fractions collected under native buffer conditions. There is fusion protein in the flowthru and the wash.

I just checked the pH of the binding, washing and eluting buffers and they were pH8.9-9.4, not pH7.4 which could explain the protein not sticking to Ni slurry.

So now, I'm thinking of several approaches:
1. Revisit native conditions and use a protease inhibitor cocktail. For sure, I am thinking of adding it to lysis buffer, but I'm wondering if I need to add it to buffers throughout the purification steps or during Tev cleavage? I do see some cleavage of fusion protein during purification. After Tev digest, I see very little of my final 8-10kDa protein even when I digest A LOT of fusion protein at 4 degrees. I am concerned the protease inhibitors will be removed during purification and desalting steps.

2. Regarding denaturing conditions, I guess I have to make the sodium phosphate buffers for His column with urea fresh just before use and pH after adding urea. This seems like an incredible pain in the neck.

3. I have a fusion protein construct with a HA-tag and I am thinking of tracking it by Western with HA tag and getting a hold of some gradient gels to improve resolution while I'm trying to work out the Tev cleavage out.

Argh! Purification of His-tagged proteins sound so simple and easy but it's so frustrating working it out.

Hi everyone
I'm doing with an His-tagged protein. Everything's ok until purification step: my protein is contaiminated with other proteins after elution. I'm using Ni-NTA agarose resin of Qiagen. They said we could use b-ME upto 20mM (I'm wordering what buffer we should add it to); or increase salt/detergent or add etOH/glycerol to wash buffer. Has anyone tried this? How about the concentration range of these components you think i should try?
Thank you.